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Abstract

Background

Despite considerable progress in the development of anticancer therapies, there is
still a high mortality rate caused by cancer relapse and metastasis. Dormant or slow-cycling
residual tumor cells are thought to be a source of tumor relapse and metastasis, and
are therefore an obstacle to therapy. In this study, we assessed the drug resistance
of tumor cells in mice, and investigated whether vaccination could promote survival.

Methods

The mouse colon carcinoma cell line CT-26 was treated with 5-fluorouracil to assess
its sensitivity to drug treatment. Mice with colon tumors were immunized with inactivated
slow-cycling CT-26 cells to estimate the efficacy of this vaccine.

Results

We identified a small population of slow-cycling tumor cells in the mouse colon carcinoma
CT-26 cell line, which was resistant to conventional chemotherapy. To inhibit tumor
recurrence and metastasis more effectively, treatments that selectively target the
slow-cycling tumor cells should be developed to complement conventional therapies.
We found that drug-treated, slow-cycling tumor cells induced a more intense immune
response in vitro. Moreover, vaccination with inactivated slow-cycling tumor cells caused a reduction
in tumor volume and prolonged the overall survival of tumor-bearing mice.

Conclusions

These findings suggest that targeting of slow-cycling tumor cells application using
immunotherapy is a possible treatment to complement traditional antitumor therapy.

Keywords:

Background

In the majority of cancer cases, mortality is caused by metastases, with only 10%
being caused by the primary tumor [1]. In many cancers, metastases and relapses may occur several years or decades after
disease remission. Disseminated tumor cells or residual treatment-resistant tumor
cells may persist in a so-called dormant state until they are stimulated into an active
cell-cycle and initiate tumor recurrence [2]. Thus, these dormant or 'slow-cycling' residual tumor cells are thought to be a source
of tumor relapse and metastasis, and are therefore an obstacle to therapy. However,
the identification and functional characterization of slow-cycling tumor cells are
still poorly understood.

It is accepted that slow-cycling tumor cells are more drug-resistant than normal tumor
cells, although direct proof of this is lacking. The suggested mechanism of the drug
resistance of slow-cycling tumor cells is that their minimal activity silences a vast
spectrum of metabolic loops targeted by anticancer drugs [3]. However, this theory is still controversial, and more research is needed.

Clinical studies have recently shown that adding immunotherapy to chemotherapy has
survival benefits compared with chemotherapy alone, and can sensitize tumors to immune-cell-mediated
killing [4]. Cancer vaccination with inactivated tumor cells is one form of immunotherapy that
is in common use. Studies that have identified slow-cycling tumor cells as the source
of tumor relapse and metastasis have also indicated their possible use in cancer vaccination.
It is likely that some proteins with distinct immunogenicity are specifically expressed
on the surface of slow-cycling tumor cells, which therefore provides opportunities
for enhanced immunotherapy.

In the present study, we investigated the tumorigenicity and drug-resistant potential
of slow-cycling tumor cells compared with normal tumor cells, and found evidence supporting
the hypothesis that slow-cycling, drug-resistant tumor cells are the source of tumor
relapse and metastasis, and are thus an obstacle to therapy. We found that, compared
with normal tumor cells, the inactivated slow-cycling, drug-resistant cells induced
greater proliferation of spleen cells and higher production of interferon (IFN)-γ
by these spleen cells in vitro. We also investigated the use of such tumor cells in cancer vaccination. We found
that vaccination using the slow-cycling, drug-resistant tumor cells induced a conspicuous
immune response in mice with colon carcinoma and remarkably prolonged the overall
survival of the animals.

Methods

Ethics

Experimental research that is reported in the manuscript have been performed with
the approval of the Animal Care and Welfare Committee of CIH-CAMS-PUMC (approval date:
20 June 2009; approval number: 20120002). All the experimental research on animals
followed the National Institutes of Health Guide for the Care and Use of Laboratory Animals (publication no. 85-23, revised 1985).

DiI staining and cell sorting

Tumor cells were stained with DiI (Dil (1,1'-dioctadecyl 3,3,3',3'-tetramethyl-indocarbocyanine
perchlorate); Invitrogen Corp., Carlsbad, CA, USA) in accordance with the protocol
for attached cells [5]. Cells were suspended at a density of 1 × 106/ml in 1640 culture medium, DiI solution was added at a concentration of 5 μl/ml and
the cell suspension was incubated at 37°C for 20 minutes. After washing with phosphate-buffered
solution (PBS) with 2% FBS, the cells were analyzed and sorted using a fluorescence-activated
cell sorting (FACS) system (Vantage SE; Becton Dickinson, Franklin Lakes, NJ, USA).

To determine the heterogeneity of tumor cells with respect to cell-cycle length in vitro, DiI-labeled cells were allowed to grow for 8 days in complete RPMI 1640 culture
medium under normal conditions, and were analyzed by flow cytometry on days 1, 3,
5, and 8. To determine the heterogeneity of tumor cells with respect to cell-cycle
length in vivo, DiI-labeled cells were injected subcutaneously into the left groin of Balb/C mice
(2 × 106 cells per mouse; four mice in total). Tumors were digested in complete RPMI medium
containing 1 mg/ml type IV collagenase and 300 U/ml DNase I (Sigma AB, Göteborg. Sweden)
incubated for 30 min at 37°C, and analyzed by flow cytometry on days 10, 15, and 25.
When sorting, the slow-cycling cells were identified as a bright positive population.

Hoechst-Pyronin Y staining and cell sorting

Cells were detached from the cell-culture flask with 0.1% trypsin, and Trypan blue-nonstaining
viable cells were counted and suspended at a density of 1 × 106/ml in DMEM culture medium. Then they were stained with the fluorescent dye Hoechst
33342 (Sigma AB) at a concentration of 5 μg/ml at 37°C for 45 minutes. At the end
of this time, 1 μg/ml of Pyronin Y (PY) was added, and cells were incubated at 37°C
for an additional 45 minutes as described previously [6]. After washing with PBS plus 2% FBS, the cells were analyzed and sorted (FACS Vantage
SE; Becton Dickinson). When sorting, cells residing in the G0/G1 peak that simultaneously
stained weakly with PY were regarded as cells in G0 phase, and these were sorted and
used for further studies [7].

Side-population analysis

Cells were detached from the cell-culture flask with 0.1% trypsin, and Trypan blue-nonstaining
viable cells were counted, and suspended at a density of 1 × 106/ml in DMEM culture medium. Then they were stained with the fluorescent dye Hoechst
33342 (Sigma AB) at a concentration of 5 μg/ml (37°C for 90 min) as described previously
[8]. After washing with PBS plus 2% FBS, the cells were incubated with 2 μg/ml propidium
iodide (PI) to exclude dead cells, then cell analysis was performed (FACS Vantage
SE; Becton Dickinson).

Tumor generation

Viable fast-cycling and slow-cycling tumor cells obtained using the DiI-based FACS,
and viable G0 and non-G0 cells obtained by Hoechst-PY-based FACS, were stained with
Trypan blue and counted. Then cells of every population were injected subcutaneously
into the left groin of Balb/C mice at a gradient dose of 5000, 1000, or 500 cells.
The mice were examined visually every day.

Chemotherapy resistant assay

To investigate the chemotherapy resistance of slow-cycling cells in vivo, DiI-labeled cells (1 × 106 per mice) were injected subcutaneously into Balb/C mice. When the tumors had grown
to 10 × 10 mm in size, 5-fluorouracil (5-FU) 40 mg/kg was injected intraperitoneally
every 3 days for a total of four injections. The vehicle control mice were injected
with PBS, using the same regimen. After the final treatment, tumors were digested
into a single-cell suspension as described above, and analyzed by flow cytometry the
next day.

To determine the chemotherapy resistance of slow-cycling cells in vitro, the same numbers of DiI-labeled cells were seeded into a cell-culture flask (day
1), and grown for 24 hours, then treated with 5-FU (day 2) at a concentration of 1.5
μg/ml. On day 3, the medium was replaced with fresh medium without 5-FU, and the cells
were grown under normal conditions for 24 hours. On day 4, 5-FU 1.5 μg/ml was added
into the medium again, and cells were grown for a further 24 hours, then on day 5,
the medium was again replaced with fresh medium without 5-FU, and cells were grown
for another 24 hours. Finally, on day 6, cells were treated with trypsin and analyzed
by flow cytometry. The control cells were treated in the same way but were never exposed
to 5-FU.

To detect the inhibition of cell proliferation by 5-FU in vitro, DiI-labeled cells of test group and control group were seeded in triplicate into
96-well culture plate at 3,000 cells/well, then challenged with 5-FU 24 hours later
in the same manner above. On day 6, 3-(4,5-dimethylthiazol-2-yl)- 2,5- diphenyltetrazolium
bromide (MTT) method was performed as described previously [9].

In vitro lymphocyte proliferation assay

Mixed lymphocyte tumor cell culture (MLTC) was used to investigate the proliferation
of spleen cells. Tumor-bearing mice were killed by broken neck and spleens were harvested.
The spleen tissue was ground and suspended in PBS, then spleen cells were isolated
using density gradient centrifugation (Ficoll-Hypaque, Haoyang Biological Manufacture,
Tianjin, China) and stored as a single-cell suspension. CT-26 cells treated with 5-FU
(FU-CT-26) or not (non-FU-CT-26) were exposed to mitomycin C (MMC) for 1.5 hours,
then these cells were seeded in triplicate at a density of 1 × 104 cells per well in 96-well culture plates, along with spleen cells (1 × 105 cells per well), and incubated with interleukin (IL)-2 (100 U/ml) for 4 days at 37°C
in a humidified 5% CO2 atmosphere. The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide)
assay was used to test the lymphocyte proliferation [9], and the results were expressed as:

where A is the experimental absorbance from the spleen plus tumor cell co-cultures,
B is the absorbance from the tumor cells alone, and C is the absorbance from the spleen
cells alone.

ELISA for the production of interferon-gamma (IFN-γ)

CT-26 cells treated with 5-FU or not were exposed to MMC for 1.5 hours, then these
cells (2 × 105 cells per well) were co-cultured separately with spleen cells (2 × 106 cells per well) from tumor-bearing mice at ratio of 1:10 in 400 μl complete RPMI 1640
containing IL-2 (100 U/ml) for 3 days. The supernatant was collected on day 4, and
the concentration of IFN-γ was analyzed using a mouse IFN-γ ELISA kit (eBioscience
Inc., San Diego, CA, USA) as described previously [10].

In vitro cytotoxic assay

Mice (three mice per group, and three groups in total) were challenged with 3 × 105 CT-26 cells injected subcutaneously into the left groin (day 0), then separately immunized
with subcutaneous injection of FU or non-FU-CT-26 cells (1 × 106) that had also been pretreated with MMC on days 3, 6, 9, 13, 18, and 25. 7 days after
the final booster. Spleen cells from the immunized mice (FU or non-FU-CT-26 groups)
were prepared as effector cells. Mice in the control group were treated in the same
way, but using PBS for injection.

4T-1, YAC-1, and FU or non-FU CT-26 cells were used as target cells. As described
previously [11], target cells were labeled with 5- (and 6-) carboxyfluorescein diacetate succinimidyl
ester (CFSE; Sigma AB) for 10 minutes at 37°C using a final concentration of 2 μmol/L.
After labeling, the cells were washed once and re-suspended in complete RPMI 1640.
Effector and target cells were mixed to a final volume of 200 μl in complete RPMI
1640, with the ratio of effector:target being 50:1. The tubes were mixed and spun
down at 120 × g for 2 minutes, then the samples were incubated at 37°C for 4 hours.
At the end of incubation time, 2.5 μg PI (Sigma AB) was added for DNA labeling of
dead cells. The samples were then incubated for 5 minutes and analyzed by flow cytometry
within 60 minutes.

Vaccination treatment in a murine colon cancer model

To establish a colon cancer model, 3 × 105 CT-26 cells were subcutaneously inoculated at the left groin of Balb/C mice on day
0 (five mice per group, and five groups in total). Then tumor cells were injected
combined with or not with granulocyte-macrophage colony-stimulating factor (GM-CSF,
1 ng per mouse) on days 3, 6, 9, 13, 18, 25. The FU and non-FU CT-26 cells used in
the earlier vaccination were pretreated with MMC and injected subcutaneously at 1
× 106 per mouse. Control mice were treated with PBS. Tumor growth was monitored every 2-3
days by palpation, and tumor size was measured in two perpendicular tumor diameters,
as described previously [12].

Statistical analysis

Statistical significance of difference between the two groups was determined by the
Student paired t-test. The Kaplan-Meier plot for survival was assessed for significance
using the log-rank test (SPSS software; version 12.0; SPSS Inc., Chicago, IL, USA).
P < 0.05 was considered significant.

Results

Tumor cells exhibit clear heterogeneity in to cell-cycle length both in vivo and in vitro

DiI is a long-term lipophilic tracer dye that can be used to trace cell division and
identify slow-cycling cells. It has several advantages compared with PKH26, such as
a simpler protocol for cell labeling, lower cytotoxicity, and higher resistance to
intercellular transfer [13]. This dye normally disappears at cell division; however, slow-cycling cells may retain
it for a long time, which allows them to be identified by flow cytometry. We found
that DiI-retaining cells disappeared when they were allowed to grow in vitro or in vivo (Figure 1A, B). A bright DiI-positive population of cells (DiIhigh) was visible after 8 days in culture (7.3% of cells; Figure 1A) or 25 days growing in vivo (0.5% of cells; Figure 1B). Tumor cells exhibited clear heterogeneity with respect to cell-cycle length both
in vivo and in vitro, with slow-cycling tumor cells (DiIhigh), cycling tumor (DiIlow) cells, and fast-cycling tumor cells (DiI-negative cells) all being present. Moreover,
slow-cycling tumor cells only comprised a small proportion of the tumor mass.

Figure 1.The DiI-tracing assay showed that tumor cells exhibited clear heterogeneity with respect
to cell-cycle length. (A) DiI-labeled CT-26 cells were cultured in complete RPMI 1640 medium on day 1, and the
percentage of DiI-retaining cells was analyzed on days 1, 3, 5, and 8 by flow cytometry.
(B) DiI-labeled CT-26 cells were injected subcutaneously into mice on day 1. On days 10,
15 and 25, tumors were excised and digested, and the percentage of DiI-retaining cells
was analyzed by flow cytometry. Experiments were repeated twice with similar results.

Slow-cycling tumor cells have the character of stem cells

Slow-cycling tumor cells are thought to be drug-resistant and the source of relapse
and metastasis. To investigate the tumorigenic potential of this cell population,
freshly sorted slow-cycling and fast-cycling tumor cells were injected subcutaneously
into Balb/C mice, and their tumor-formation ability was assessed.

A bright DiI-retaining population of cells was selected as slow-cycling cells, and
a population of DiI-negative cells was selected as fast-cycling cells. After injection
of 5,000 DiIhigh or DiI-negative cells, all five mice that received the DiIhigh cells developed tumors, whereas only three of five mice that received DiI-negative
cells developed tumors. Similarly, when the number of cells injected decreased to
1000, tumors formed in three of five mice that received DiIhigh cells, whereas tumors were only seen in one of five mice injected with the DiI-negative
cells. After injection of 500 DiIhigh cells, only one of five mice had established tumors, whereas no tumors were established
from DiI-negative cells (Table 1).

Analogous results were seen in cells sorted by Hoechst-PY stain-based FACS. There
are many dyes available for determining G0/G1 versus S and G2/M phases based on the
DNA content measured by flow cytometry. However, a DNA dye is not able to distinguish
cells residing in G0 or G1 phases. This can be achieved by quantifying RNA content
(which increases during G1 and remains high during mitosis) using PY in conjunction
with the DNA dye Hoeschst 33342 [14]. Tumor cells residing in the G0/G1 peak and simultaneously weakly stained by PY were
sorted as G0 cells (P2; Figure 2A), whereas tumor cells residing in S phase were regarded as non-G0 cells (P3; Figure
2A). The results of the tumor-generation assay were similar to those of the assay mentioned
in the previous paragraph. Tumors formed in all five mice injected with 5,000 G0 cells,
compared with two of five mice injected with 5,000 non-G0 cells (Table 2); in all five mice injected with 1,000 G0 cells, compared with only one of five mice
injected with 1,000 non-G0 cells; and in three of five mice injected with 500 G0 cells,
compared with none of the mice injected with non-G0 cells. All these results indicated
increased tumorigenicity of slow-cycling tumor cells.

Figure 2.Most cells in the G0 phase sorted by fluorescence-activated cell sorting (FACS) also
resided in the same region as side-population cells. (A) Hoechst-Pyronin Y (PY)-based FACS. The left and middle panels separately represent
the Hoechst-staining control group and PY-staining control group. In the Hoechst-PY-staining
group (right), the P2 population represents the cells residing in G0 phase, and P3,
cells in S phase. (B) G0 cells sorted by Hoechst-PY-staining-based FACS (left panel) were measured by side-population
analysis, and the result is shown in the middle panel. The right panel represents
the classic side-population analysis of CT-26 cells. The middle and right panels show
that most of the cells in G0 phase also resided in the region of the side-population
cells. Experiments were repeated three times with similar results.

Until recently, side-population analysis has been one of the accepted methods for
identifying cancer stem cells (CSCs) [15]. We investigated the percentage of side populations in CT-26 cells (Figure 2B, right). We then analyzed the number of G0 cells sorted by Hoechst-PY staining-based
FACS (Figure 2B, left) using side-population analysis (Figure 2B, middle). G0 cells resided in almost the same part of the coordinate axis (Figure
2B, middle) as the side-population cells (Figure 2B, right), and they had a similar proportion of tumor cells (2.6% versus 2.4%; Figure
2B, middle and right). These findings and the increased tumorigenicity of slow-cycling
tumor cells indicated that they might have the character of CSCs.

We found that although slow-cycling tumor cells exhibited a higher tumorigenic potential,
the average number of days of tumor generation was prolonged compared with that of
fast-cycling cells (Table 1, Table 2).

Slow-cycling tumor cells are more resistant to chemotherapy, both in vivo and in vitro

Slow-cycling tumor cells have always been predicted to be resistant to traditional
chemotherapy. We treated CT-26 cells with 5-FU in vivo and in vitro to investigate the sensitivity of slow-cycling cells and normal tumor cells.

Tumor volume in mice that received chemotherapy decreased markedly compared with that
in PBS-treated mice (Figure 3B). However, after four treatments with 5-FU, the percentage of DiI-retaining slow-cycling
cells increased significantly, being 12% in tumors from mice that received chemotherapy
compared with 2.8% in tumors from mice treated with PBS (Figure 3A, C).

Figure 3.Chemotherapy-induced enrichment of DiI-retaining cells in vivo. (A) DiI-retaining tumor cells were enriched in vivo after treatment with 5-fluorouracil (FU). Negative: flow cytometry of CT-26 cells
that were not stained with DiI: PBS-treated: flow cytometry of cells from tumor treated
with phosphate-buffered saline (PBS); 5-FU-treated: flow cytometry of cells from tumor
treated with 5-FU. (B) Tumor volume in mice after chemotherapy compared with the PBS-treated group (*P < 0.05, t-test). (C) Percentage of DiI-positive cells in tumors with or without chemotherapy (**P < 0.01, t-test). Error bars represent the standard deviation. DiI-labeled CT-26 cells were
injected subcutaneously into Balb/C mice on day 1. When tumors grew to 10 × 10 mm,
5-FU 40 mg/kg was injected intraperitoneally every 3 days for a total of four injections.
Vehicle-treated control mice were exposed to the same regimen, but injected only with
PBS. Tumor volume was measured on days 7, 14, and 21. After the final treatment, tumors
were digested and analyzed by flow cytometry on the next day. Experiments were repeated
twice with similar results.

Similarly, in vitro, 5-FU treatment obviously inhibited the proliferation of tumor cells, as indicated
by the absorbance of MTT (Figure 4B). Tumor cells after chemotherapy were analyzed by flow cytometry; most cells were
DiI-retaining slow-cycling cells (98.2%), while the percentage of DiL-positive cells
in the untreated group was only 2.8% (Figure 4A, C). These findings were consistent with the previous prediction that slow-cycling tumor
cells are an obstacle to traditional chemotherapy.

Figure 4.Chemotherapy-induced enrichment of DiI-retaining cells in vitro. (A) DiI-retaining tumor cells were enriched in vitro after treatment with 5-fluorouracil (FU). DiI-labeled CT-26 cells were cultured in
complete RPMI 1640 medium and exposed to 5-FU on day 2. The old medium was changed
for fresh medium without 5-FU on day 3, then 5-FU was added again on day 4. Cells
were harvested on day 6 and analyzed by flow cytometry. The control group was cultured
without 5-FU. (B) Persistence of cells after 5-FU treatment. The same numbers of CT-26 cells were seeded
into 96-microwell plates. The 5-FU-treated group was treated as described above, and
the untreated group was cultured without 5-FU. An MTT assay was performed 5 days later.
(**P < 0.01, t-test) (C) Percentage of DiI-positive cells in vitro after 5-FU treatment. (**P < 0.01, t-test). Experiments were repeated twice times with similar results. Error bars represent
the SD.

5-fluorouracil-treated CT-26 cells induce increased proliferation of and interferon-γ
production by spleen cells in vitro

To investigate the potential of inducing proliferation of and IFN-γ production by
spleen cells in vitro, FU and non-FU CT-26 cells were treated with MMC and then co-cultured with freshly
isolated spleen cells in medium containing IL-2. After 3 or 4 days culture, an MTT
assay was performed to analyze the proliferation rate. To investigate the efficacy
of MMC, the proliferation of MMC-treated FU-CT-26 and non-FU-CT-26 cells was analyzed.
No proliferation was seen in either group of tumor cells after treatment with MMC,
which indicates that MMC could be used to inactivate both FU and non-FU CT-26 cells
(Figure 5B). The FU-CT-26 cells had a higher proliferation index than the non-FU-CT-26 cells
(2.11 versus. 1.70; Figure 5B).

Figure 5.5-fluorouracil (FU)-treated CT-26 cells induced increased proliferation and interferon
(IFN)-γ production by spleen cells. (A) Proliferation curve of FU-treated and non-FU-treated CT-26 cells after treatment with
mitomycin C (MMC). No proliferation was seen with either of the tumor cells, which
confirmed the efficacy of MMC. (B) Spleen cells from tumor-bearing mice had significantly higher proliferation after
co-culture with FU-CT-26 cells compared with non-FU-CT-26 cells, at a responder:stimulator
(R:S) ratio of 10:1. (*P < 0.05, t-test) (C) IFN-γ production by spleen cells after co-culture with 5FU-treated CT-26 cells compared
with CT-26 cells. (*P < 0.05, t-test). Error bars represent the standard deviation. CON: control spleen cells cultured
without tumor cells. Mixed lymphocyte tumor cell culture (MLTC) was performed to investigate
the proliferation of and IFN-γ production by spleen cells. Spleen cells were harvested
from tumor-bearing Balb/C mice. The same numbers of FU-CT-26 and non-FU-CT-26 cells
were seeded into 96-microwell plates. Spleen cells were added at an R:S ratio of 10:1
on day 1, and proliferation of spleen cells was analyzed by MTT assay on day 4. As
for IFN-γ production, the same numbers of FU-CT-26 and non-FU-CT-26 cells were seeded
into 24-well plates. Spleen cells were added at an R:S ratio of 10:1. The supernatant
was collected on day 4, and IFN-γ concentration was analyzed by ELISA.

Culture supernatant was collected and used for IFN-γ analysis by ELISA. FU-CT-26 cells
induced more IFN-γ production in spleen cells compared with CT-26 cells (Figure 5C). The concentrations of IFN-γ in the supernatant of the FU-CT-26 plus spleen cell
mixture was 1602 ± 55, that of the non-FU-CT-26 plus spleen cells was 750 ± 24, and
that of the spleen cells alone was 54 ± 11 pg/ml, respectively.

The preceding results provided strong information that slow-cycling tumor cells may
be a better immunogen that could induce an intense antitumor response. To investigate
whether immunization with inactivated FU-CT-26 cells could obtain a better therapeutic
effect in vivo, we established a subcutaneous CT-26 mouse model and immunized them separately with
inactivated CT-26 and FU-CT-26 combined with GM-CSF. Tumors in mice treated with FU-CT-26
or non-FU-CT-26 cells plus GM-CSF, or non-FU-CT-26 cells alone were all clearly reduced
compared with those in mice treated with PBS (control group). Immunization with the
FU-CT-26 cells plus GM-CSF produced the best therapeutic effect (Figure 6A). Treatment with FU-CT-26 cells plus GM-CSF prolonged survival of tumor-bearing mice
(Figure 6B). Both the mice treated with PBS and the mice treated with the non-FU-CT-26 cells
all died, with a median survival of 32.8 and 40.2 days, respectively. The mice immunized
with FU-CT-26 and non-FU-CT-26 cells plus GM-CSF had comparable results, with median
survival of 50.7 and 48.5 days, respectively. When survival was monitored for up to
80 days after inoculation, one mouse in each group was still alive. However, treatment
with FU- CT-26 cells plus GM-CSF exhibited the best outcome, with a median survival
of 61.5 days, and a 60% survival rate (three of five mice) when survival was monitored
up to 80 days after inoculation.

We rechallenged the mice that survived after immunotherapy (one treated with FU-CT-26
cells alone, one with non-FU-CT-26 cells plus GM-CSF, and three with FU-CT-26 cells
plus GM-CSF) with 106 non-FU-CT-26 cells. None of these five surviving mice had established tumors, which
indicated that immunotherapy with inactivated FU-CT-26 and non-FU-CT-26 cells induced
a specific memory immune response in vivo.

Immunization with inactivated FU-CT-26 cells showed a clear therapeutic effect in vivo, therefore, we investigated the mechanism involved. First, we investigated the sensitivity
of slow-cycling tumor cells to killing by cytotoxic T lymphocytes (CTLs). Spleen cells
from tumor-bearing mice were isolated, and their cytotoxicity against FU-CT-26 and
non-FU-CT-26 cells was analyzed in vitro. CTLs showed 11.85% cytotoxicity against FU-CT-26 cells compared with 21.67% cytotoxicity
against non-FU-CT-26 cells (Figure 7A). These data indicate that slow-cycling, drug-resistant tumor cells were also resistant
to cytotoxic killing, and this coincided with the clinical data.

Upregulation of major histocompatibility complex and co-stimulatory molecules on the
surface of fluorouracil-treated CT-26 cells

Using flow cytometry, we analyzed the expression of MHC class I and II molecules and
co-stimulatory molecules on the surface of FU and non-FU CT-26 cells. Although the
expression of MHC class I molecules was comparable in both FU and non-FU cells (92.7
± 2.76% versus 99.1 ± 1.27%), the average fluorescence intensity of MHC class I expressed
by the FU-CT-26 cells was much lower (13.6 ± 0.21 versus 38.9 ± 2.34). However, compared
with the non-FU-CT-26 cells, the expression level and average fluorescence intensity
of MHC class II molecules and of the co-stimulatory molecules CD80 and CD86 on the
surface of FU-CT-26 cells were all clearly upregulated (Table 3), although the expression level of MHC class II molecules and CD86 was not high.

Table 3. Expression levels and fluorescence intensity of MHC and co-stimulatory molecules on
the surface of 5-FU-treated cellsa, b.

To confirm this finding, we analyzed the expression of the molecules on the surface
of 5-FU-treated 4T-1 and TC-1 or untreated 4T-1 and TC-1 tumor cells by flow cytometry,
and a similar tendency was seen (Table 3). These findings indicate that increased expression of MHC class II molecules and
the co-stimulatory molecules CD80 and CD86 may be the reason why drug-treated tumor
cells can induce a more significant immune response, and lower expression of MHC class
I molecules could be one reason for the resistance of slow-cycling tumor cells to
cytotoxic killing.

Discussion

Tumor dormancy has been recognized for many years as a clinical phenomenon in several
types of cancer. Clinicians and experimental biologists have used the term 'dormancy'
to describe the hypothetical state of cancer cells lying in wait for some time after
treatment of the primary tumor, before the tumor's subsequent growth and clinical
recurrence [2,16]. Tumors in dormancy are mainly constructed of quiescent or slow-cycling tumor cells.
Quiescent tumor cells can be detected in the marrow of many patients in the tumor-remission
phase, and these patients often develop tumor relapse or metastasis [17-19]. However, there is insufficient evidence to prove that these cells are the origin
of tumor relapse. Thus, more research into the identification and biologic character
of quiescent or slow-cycling tumor cells is needed.

In the present study, we used a membrane-bound dye, DiI, to identify slow-cycling
cancer cells in vitro and in vivo. Our data directly confirm the existence of quiescent cells in growing colon tumor,
and this cell population comprised only a small proportion of the tumor mass. Compared
with other label-retention methods, DiI is simpler to use and yields easy identification
of quiescent, label-retaining cells. However, it is important to note that the best
time for analysis will differ depending on the type of tumor, because of the distinct
proliferation cycle of different cells.

Many human cancers contain CSCs that are responsible for initiating and maintaining
tumor growth and resistance to therapy [20-23]. The quiescent state seems to be necessary for preserving self-renewal of stem cells
[24], and is a crucial factor in resistance to chemotherapy and targeted therapies [25-27]. In the present study, we used a tumor-forming assay to show the self-renewing potential
of slow-cycling tumor cells in vivo. Simultaneous side-population analysis of the cell line indicated that CSCs were
enriched in the slow-cycling population. It was particularly interesting that, although
more transplanted tumors were seen in mice injected with slow-cycling tumor cells,
the average tumor-forming time was longer than with the fast-cycling cells (Table
1, Table 2). This may because slow-cycling tumor cells take a long time to exit the quiescent
state, and then expand and differentiate in response to stress. This finding indicates
that, if the mechanism that causes recycling of quiescent cells could be elucidated
and the crucial point of the pathway inhibited, this recycling could be inhibited,
preventing tumor relapse and metastasis. Moreover, we found that, although the number
of tumor cells and the volume of the tumor were reduced by drug treatment, the remnant
was composed of drug-resistant, slow-cycling cells. These results provide evidence
that slow-cycling tumor cells are resistant to traditional chemotherapy and are responsible
for initiating tumor relapse and metastasis.

Conventional chemotherapy optimally targets highly proliferative tumor cells, and
the existence of drug-resistant, slow-cycling tumor cells limits improvements in recurrence-free
and overall survival rates. In this study, we found that drug-resistant tumor cells
are mostly slow-cycling, and this population increased the proliferation of and IFN-γ
production by spleen cells in vitro. Moreover, our in vivo experiments showed that, compared with normal tumor cells, vaccination with slow-cycling
tumor cells generated a more effective immune response and prolonged the overall survival
of tumor-bearing mice.

Although the slow-cycling population was more resistant to CTL cytotoxicity than the
conventional tumor cells, this population could induce a more intense immune response,
as shown by the enhanced cytotoxicity of spleen cells from mice immunized with slow-cycling
tumor cells. More importantly, we found that these slow-cycling cells expressed a
lower level of MHC class I molecules, but a higher level of class II, as well as a
higher level of the co-stimulatory molecules CD80 and CD86, compared with conventional
tumor cells. We speculate that the low expression of MHC class I molecules may have
caused the resistance to killing by CTLs, whereas the upregulation of MHC class II
and co-stimulatory molecules may be one reason for the increased induction of the
immune response.

However, more questions remain about the mechanisms underlying the apparently superior
outcomes from vaccination with slow-cycling tumor cells. For example, is there any
difference between slow-cycling tumor cell antigens and conventional tumor lysates
in inducing effector cell differentiation and memory T-cell generation? Further studies
into different aspects of these tumor cells are needed. For instance, differences
in gene expression between slow-cycling and conventional tumor cells have been analyzed
by gene chip technology, and we have now found a series of overexpressed genes in
slow dividing cells. One of these antigens, which has been reported to be a testicular
cancer antigen, has particularly attracted our attention. However, further research
into this gene and its related protein is needed.

The results of the present study all indicate that slow-cycling tumor cells are a
better source of antigens for cancer immunization than conventional tumor cells. To
date, the primary treatment for eliminating slow-cycling tumor cells is to induce
them to enter the cell cycle and then kill them using traditional methods [2,28]. However, immunotherapy, as performed in our study, could selectively target the
only slow-cycling tumor cells, resulting in elimination of the source of tumor recurrence
and metastasis. Compared with conventional treatment, this technique could effectively
reduce the risk of tumor recurrence and metastasis. Although several studies have
shown that vaccination using stem-cell antigens induces a more effective immune response
against prostate, brain, and ovarian cancers [29-31], there is controversy regarding the identification and isolation of CSCs in different
tumors. Our results indicate that slow-cycling tumor cells could enrich CSCs, and
the process we used to harvest slow-cycling tumor cells is easier to perform. Thus,
the clinical application of this immunotherapy shows good prospects.

Conclusions

In this study, we showed that slow-cycling tumor cells induced an antitumor immune
response, especially of tumor-specific CTLs, with enhanced killing of drug-resistant
tumor cells, and vaccination with slow-cycling tumor cells could prolong the overall
survival of tumor-bearing mice. Our data also indicated that this treatment not only
kills normal tumor cells, but also selectively targets the slow-cycling tumor cells,
thus reducing the risk of cancer metastasis and relapse. Moreover, this vaccine has
excellent histocompatibility, because slow-cycling tumor cells are isolated from the
tissues of the recipient; thus, no severe side-effects should occur. To our knowledge,
this is the first study of its kind. All our findings suggest that immunotherapy with
inactivated slow-cycling tumor cells is a possible strategy to complement traditional
cancer treatment.

Competing interests

The authors declare that they have no competing interests.

Authors' contributions

SZ conceived of the study; SZ, QS and YZ participated in the design and coordination
of the study; QS carried out the experiments, analyzed the data, and wrote the manuscript;
FW performed the acquisition of data; CZ, DW, and WM carried out parts of the experiments
and contributed to the guidance of experiments; and YH Z read the manuscript and revised
it for important intellectual content. All authors have read and approved the final
manuscript.

Acknowledgements

We thank Dr Shengdian Wang and Youhui Zhang for their reading and suggestions in preparation
of the manuscript, and Shiliang Ma and Jianming Liang for their excellent work with
the flow cytometry. This study was supported by State Key Development Program of Basic
Research of China (Item Number: 2012CB917100) and Research Fund from Cancer Institute,
Chinese Academy of Medical Sciences.